One major area of investigation of our lab was the actomyosin cytoskeleton. Cells use the actomyosin cytoskeleton to modulate their size and shape, to crawl through different substrates, to extend out from cell masses, and to adapt to different tissue-specific environments. These processes are critical for the morphogenetic pathways underlying tissue regeneration and remodeling, as well as tissue invasion in cancer, so understanding actomyosin organization and dynamics could have multi-fold applications. To investigate the actomyosin cytoskeleton's role, we employed quantitative imaging, modeling and biophysical approaches. This included live cell 3D structured illumination microscopy; force spectrum microscopy; and, lattice light sheet microscopy. These investigations led to significant new mechanistic insights into how the actin cytoskeleton controls cell-cell interactions. In collaborative work with other labs, we were also able to demonstrate a role for actin contractile forces in mediating diffusive-like, non-thermal motion in the cytoplasm that causes intracellular movement of small and large cell components, as well as the motion of primary cilia. Related to actomyosin's role at cell-cell contacts, we used cytotoxic T lymphocytes (CTLs) as a model system. CTLs kill target cells by secreting granules containing perforin and granzymes into the immunological synapse (the site of contact formed between the CTL and target cell). We used spinning disk confocal and lattice light sheet microscopy to obtain unprecedented spatial and temporal resolution of the actin cytoskeleton and its role in both facilitating and limiting CTL secretion. We saw dynamic lamellapodial projections and a rearward flow of actin in migrating CTLs as these cells engaged a target cell. The synapse then formed in two stages: concentration of T cell receptors (TCRs) in the PM through lateral translocation (1 min), followed by vesicular delivery of intracellular TCRs as the centrosome reached the synapse (6 min). Prior to synapse formation, a continuous actin meshwork underlies the entire plasma membrane; however, local clearing occurred as both the centrosome and granules docked. After several vesicles fused, the actin meshwork reappeared and secretion stopped. Actin clearance and reappearance correlated with the loss and gain of PI(4,5)P2 in the contact zone. We concluded that the CTL contact zone is like a radially symmetric leading edge, with the distal region of protrusive actin polymerization being analogous to the lamellipodium, and the more central region, enriched in integrins and myosin IIA, analogous to the lamellum. The spatial-temporal regulation of actin in the contact zone serves to coordinate TCR docking and the timing of granule secretion. We also examined the role of the actin cytoskeleton in regulating overall motion within the cytoplasm. We reasoned that ensemble forces from actomyosin activity could have a large effect on global motion within the cytoplasm, making these forces a critical readout of the dynamic state of the cell. To quantify these forces and test how they control the motion of cytoplasmic components, we collaborated with physicist Dr. David Weitz, who devised a new methodology called force-spectrum-microscopy (FSM) to quantify force fluctuations within the cytoplasm. The technique combines measurements of the random motion of probe particles with independent micromechanical measurements of the cytoplasm. Increased cytoplasmic force fluctuations substantially enhanced intracellular movement of small and large components, including organelles. Cytoplasmic force fluctuations varied between cell types and were three times larger in malignant cells than in normal cells. Separately, in close collaboration with Christoph Schmidt, we found that force generation by the actin cytoskeleton surrounding the basal body causes previously undocumented active primary cilia movements, which could be important for tuning and calibrating ciliary sensory functions. The results of these studies reveal that actomyosin dynamics are a critical readout of proper cell health that have major effects on diverse cellular functions. A second major theme in the lab was related to mitochondrial dynamics, including mitochondrial interactions with other organelles within the cell. How these interactions impact the physiology of cells is not clear. To study mammalian cell adaptation to nutrient starvation, we examined the interplay between mitochondria fusion dynamics, autophagy, fatty acid (FA) trafficking and LDs. Given that cells appear to adapt to nutrient starvation by shifting their metabolism from reliance on glucose metabolism to utilization of mitochondrial FA oxidation, we developed an assay to investigate how FAs become mobilized and delivered to mitochondria. Using a pulse-chase labeling method to visualize movement of FAs in live cells, we demonstrated that starved cells primarily use LDs as a conduit to supply mitochondria with FAs for &#946;-oxidation. This occurred through lipase-mediated FA mobilization from mitochondria associated LDs, rather than autophagy (contrary to the pathway used by yeast cells). Autophagy contributed to the altered metabolic scheme by recovering lipids from degraded organelles, which could be used to refill LDs. Notably, mitochondrial tubulation was essential for distribution of FAs throughout the mitochondria network. Defects in mitochondrial fusion led to massive alterations in cellular FA routing. Not only did non-metabolized FAs get redirected to and stored in LDs, they were excessively expulsed from cells. Given that FAs are toxic at high levels and serve as signaling molecules at low levels, these results suggest defects in mitochondria dynamics and FA trafficking pathways may underlie the pathologies of many metabolic diseases such as diabetes and obesity. In a different mitochondrial-related project, we uncovered new machinery regulating mitochondrial fission. Seminal work from others showed that before mitochondrial division by Drp1, ER tubules encircle and constrict mitochondria. Constriction results from actin polymerization controlled by the ER-localized formin protein, INF2. How ER tubules recognize mitochondria and facilitate fission, however, is unclear. In investigating this question, we discovered a novel mitochondria-localized actin-nucleating protein, Spire1C, which interacts with INF2 on the ER. Cooperation between Spire1C and INF2 enhanced actin assembly selectively at ER/mitochondria intersections, facilitating mitochondrial constriction. We are proposing, therefore, that during mitochondrial division a Spire1C-INF2 interaction tethers the ER to mitochondria and mediates actin polymerization, resulting in mitochondrial constriction.